Characterization of Collagen Abnormalities in Abdominal Aortic Aneurysms
THESIS
Presented in Partial Fulfillment of the Requirements for Honors Research Distinction in Biomedical Engineering at The Ohio State University
By
Anna Debski
Undergraduate Program in Materials Science and Engineering
The Ohio State University
2020
Thesis Defense Committee:
Professor Gunjan Agarwal, Adviser
Professor Heather Powell
i
Copyrighted by
Anna Debski
2020
ii
Abstract
Abdominal aortic aneurysm (AAA) is a life-threatening vascular disease, characterized by
abnormal dilatation of the aorta and remodeling of the elastin and collagen components of the
extracellular matrix (ECM). While elastin remodeling is well characterized and is understood to
dictate the vessel wall’s architecture and stability, little is known about microstructural changes
regarding collagen remodeling.
This study focuses on identifying characteristics of collagen remodeling in AAA which
can eventually be applied to understand pathogenesis and therapeutic techniques in AAA. Studies
were conducted on aortic tissue obtained from mouse models of AAA and clinical AAA tissue
excised at the time of vascular surgery. Non-AAA control aortic samples from each species were
also utilized. Atomic force microscopy (AFM), a high-resolution nanoscale imaging technique was
used to observe the sample topography and characterize collagen fibrils. Tissues were also stained
using collagen hybridizing peptide (CHP) and analyzed using fluorescent microscopy and second
harmonic generation (SHG) microscopy to locate regions of healthy and degraded collagen.
Our results indicate that a significant fraction of collagen fibrils in AAA tissues departed
from their native structure. These ‘abnormal’ fibrils had unresolvable D-period bands and a
wavier, appearance. AFM analysis revealed a significant reduction in the depth of D-periods in
these abnormal fibrils. Additionally, regions of abnormal collagen were located within the
remodeled areas of AAA tissue and were distinct from healthy collagen regions as ascertained
using CHP staining and SHG. Quantifying the amount of degraded collagen in AAA tissue and
understanding the causes of abnormal collagen remodeling can provide novel insights into the
ECM remodeling process in AAA and other cardiovascular diseases.
iii
Acknowledgements
I would like to thank my research advisor Dr. Gunjan Agarwal for her continued support and
guidance in my research and academic endeavors. Without her I would not have had the chance
to experience the opportunities I had throughout my undergraduate career. Dr. Agarwal has
provided me with great mentorship and has encouraged me throughout this entire project. She
has helped me become a better student, better researcher and better engineer.
I would also like to thank Blain Jones for his time and efforts in transitioning me to this project
and his continued guidance as it progressed. Without his help, the project would not have been
completed.
I would also like to thank Dr. Powell for serving on my defense thesis. The classes she taught
have provided me with extremely valuable lessons, both related and unrelated to academia.
Throughout my time at the university, she has helped me make several tough decisions and her
continued encouragement has motivated me throughout my journey.
Lastly, I would like to thank my fellow lab mates in Dr. Agarwal’s lab. Their help with my
project has enabled its completion. I have learned throughout my time in the ab from them and
will continue to employ what they have taught me in my future experiences. It has been great
working with all of them.
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Vita
2016…………………...…….Bridgewater Raritan Regional High School
2016 to present…………...…B.S. Materials Science and Engineering, The Ohio State University
Fields of Study
Major Field: Materials Science and Engineering
Publications
Jones, B. Tonniges, J. R., Debski, A., Albert, B., Yeung, D. A., Gadde, N., Mahajan, A., Calomeni,
E. P., Go, M. R.,Hans, C. P., Agarwal, G. Collagen fibril abnormalities in human and mice
abdominal aortic aneurysm, Acta Biomaterialia. 2020.
Hogrebe, N., Reinhardt, J.W., Tram, Debski, A. C., Agarwal, G, N., Reilly, M.A., Gooch, K.J. Independent
control of matrix adhesiveness and stiffness within a 3D self-assembling peptide hydrogel. Acta
Biomateriala, 2018;70:110-119.
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Table of Contents
Abstract………………………………………………………………………………….……..…ii
Acknowledgements……………………………………………………………………….……....iv
Vita……………………………………………………………………………………….………..v
Fields of Study………………….………………………………………………………….……...v
Publications….………………….………………………………………………………….……...v
Table of Contents………………………………………….……………………………..……….vi
List of Figures……………………………………….………………………….………….…….vii
List of Tables……………………………………………………………………………..….….viii
Introduction……………………………………………………………..………………….……...1
Background………………..………………………………………………………………1
Research Goals……………………………………………………………………………3
Materials and Methods…………………………………………………………………………….3
Mice……………………………………………………………………………………….3
Clinical Tissue…………………………………………………………………………….4
Atomic Force Microscopy (AFM)…………………………………………………….….4
Collagen Hybridizing Peptide…………………………………………………………….5
Statistical Analysis………………………………………………………………………..6
Results and Discussion……………………………………………………………………………6
Conclusion……………………………………………………………………………………….16
References……………………………………………………………………………………….17
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List of Figures
Figure 1: Verification of abnormal collagen fibrils in AAA using AFM…………………..……7
Figure 2: Additional examples of abnormal collagen fibrils…………………………………….8
Figure 3: Characterization of depth of D-periods in murine aortas ……………………………10
Figure 4: Characterization of depth of D-periods in human aortas………………………..……11
Figure 5: Fluorescent imaging of CHP signal in murine aortas………………………………...14
Figure 6: SHG analysis of denatured collagen………..…………………………….…………..15
vii
List of Tables
Table 1: Summary of measurements of D-depth of collagen fibrils.………………………….....12
1
Introduction
Background
Abdominal aortic aneurysm (AAA) is a vascular disease characterized by abnormal
dilations in the walls of a vessel1, which can expand, weaken and rupture, leading to fatal internal
bleeding1. It is a major contributor to mortality in the United States5. AAA is characterized by
remodeling of elastin and collagen components of the extracellular matrix (ECM)6. The current
and only technique used clinically to detect AAA is through monitoring the size (diameter) of the
aorta2. Aortas with a diameter >5.5 cm are recommended for surgical repair while aortas with a
smaller diameter are not. However, this is not a foolproof technique as some small-diameter aortas
continue to grow and rupture, while some large-diameter aortas remain stable3,4. The
unpredictability of AAA has motivated efforts to better characterize AAA, which could potentially
lead to clearer identification techniques and creation of therapeutic strategies for AAA.
The primary constituents of the ECM in the vessel wall are elastin and collagen. The
quantity, organization and structure of collagen and elastin fibers control the characteristics of the
vascular tissue7. The remodeling of collagen and elastin that occurs in AAA has an impact on the
AAA growth and aneurysm rupture1. Elastin remodeling is well understood in AAA and is
characterized by elastin disruption, increased crosslinking in the medial layer and a higher degree
of degradation than in healthy tissue8. The extent of the elastin remodeling has also been identified
through ultrastructural imaging9. Initial remodeling is characterized by formation of porous elastin
by the generation of holes throughout the tissue9. More intense remodeling is marked by clumps
of elastin fibers and severe remodeling is identified by electron-translucent elastin5. Degraded
2
elastin is also understood to serve as site(s) for deposition of calcium phosphates affecting the
vessel wall compliance10. Similar characteristics of elastin remodeling have also been identified in
other vascular diseases, indicating common features across a range of diseases11.
While much is understood about elastin remodeling, little is known about collagen
remodeling in AAA12. Collagen abnormalities have previously been identified at the fiber level.
Scanning electron microscopy and polarized light microscopy have uncovered that adventitial
collagen fibers are less undulating than control tissue samples13. Another technique, second
harmonic generation microscopy also revealed a less prominent waviness of collagen fibers in
AAA in both human and mouse models14,15. However, much less is understood about the
abnormalities in the sub-fiber level of collagen in AAA and other vascular diseases. The sub-fiber
structure of collagen i.e. the collagen fibril is instrumental to macromolecular ECM remodeling
and a determinant of the functional properties of the vessel wall and the aneurysm.
Structural changes in the collagen fibrils are well characterized in diseases characterized
by collagen mutations such as ostoegenica imperfecta and Ehlers Danlos syndrome16. However
very little is understood about structural changes in collagen fibrils in diseases without any
underlying mutation. It is interesting to note that a few reports exist on structural changes in
collagen fibrils in the dermal connective tissue of patients with vascular diseases9. These
abnormalities include changes in fibril diameter or spacing, irregular cross-sections or lack of D-
periodic banding17,18. The evidence of collagen abnormalities in dermal tissues led us to
hypothesize that collagen fibril abnormalities would be present in the vascular tissue in AAA.
3
Research Goals
The goal of this study was to identify abnormalities in collagen fibrils in AAA in order to
better understand the pathology of the remodeling process. AAA tissue across two different species
(mice and human) were examined using ultrastructural atomic force microscopy (AFM). Tissues
were also examined for collagen degradation by using a novel reagent, namely a collagen
hybridizing peptide (CHP). Fluorescent imaging and second harmonic generation (SHG)
microscopy were used to identify regions of intact and degraded collagen.
Materials and Methods
Mice: All mice were handled accordingly following a protocol approved by the Animal Care and
Use Committee (ACUC) at the University of Missouri (Columbia, MO). All animal research
experiments adhered to guidelines established at the NIH (Guide for the Care and Use of
Laboratory Animals). Only male mice were used for the study due to low occurrence of AngII-
induced AAA in female mice. AngII was used to induce AAA. At 8-10 weeks, male ApoE-/-
knockout mice were either infused with saline or with 1000 ng/min per kg of AngII from a mini-
osmotic pump that was subcutaneously inserted. For implantation of the pumps, mice were
anesthetized in a closed chamber with 3% isoflurane in oxygen for 2-5 minutes. Each mouse was
then removed and taped to a heated procedure board (35-37°C) with 1.0-1.5% isofluorane
administered via nosecone. Proceeding 28 days, the mice were euthanized with 100 mg/kg of
ketamine and 20 mg/kg of xylazine, and the abdominal aorta was then harvested. The AAA was
confirmed as dilated through histological evaluations.
4
Clinical Tissue: Human AAA tissue was obtained following an approved Institutional Review
Board (IRB) protocol from The Ohio State University from patients undergoing surgery for the
removal/treatment of AAA. All tissue analysis was performed following guidelines outlined in the
Declaration of Helsinki and tissue was collected after obtaining a signed informed consent form
from eligible patients. Patient were being treated at the Ross Heart Hospital between the times of
January 2017 and February 2020. Patients were diagnosed with AAA if aorta diameters were
greater than 5.5 cm. The aorta sizes were measured using CT scans. Patients who were diagnosed
with having AAA were recommended to surgery based on the guidelines presented by the Society
for Vascular Surgery. Tissue collected was obtained from patients ranging from 65 to 75 years in
age. Non-aneurysmal control human tissue samples were harvested from the infrarenal section of
the aorta during autopsies performed within 24 hours of the time of death at the Detroit Coroner’s
Office. Tissue use was approved by the Institutional Review Board of Wayne University, Detroit,
MI and analysis was performed following guidelines outlined in the Declaration of Helsinki.
During surgery, an approximate 3 x 3 cm segment was removed from the anterior aortic sac of the
vessel wall and used for analysis.
Atomic force microscopy: Tissue segments obtained previously obtained from AngII-infused
mice, saline-infused mice, clinical AAA human samples and non-aneurysmal human samples were
dissected and fixed in 2% glutaraldehyde. The tissue samples were then embedded in Optimal
Cutting Temperature (OCT) compound and flash frozen in liquid nitrogen. Sections were stored
at -80oC for future use. When used for AFM analysis of topography, the OCT embedded tissues
were cryosectioned into 5 μm thick sections and mounted onto poly-lysine coated glass cover slips,
washed with purified water, and air dried. The sections were then imaged using with a Multimode
5
AFM equipped with a JV scanner and a Nanoscope Illa Controller. Samples were imaged using
tapping mode in ambient air with NSC15 cantilevers (Mikromasch). AFM height and amplitude
images were obtained at 512 lines per scan direction and a scan speed of ~ 2 Hz. The length and
depth of D-periods in collagen fibrils was ascertained by section analysis of the collagen fibrils in
AFM height images using the Nanoscope software.
Collagen hybridizing peptide (CHP) staining: Aorta segments obtained from mouse models
were used unfixed. Human AAA tissue was fixed with 4% paraformaldehyde. The tissue samples
were then embedded in Optimal Cutting Temperature (OCT) compound and flash frozen in liquid
nitrogen. Sections were stored at -80oC for future use. For staining, the OCT embedded tissues
were cryosectioned into 5 μm thick sections and mounted in triplicates onto microscope slides
(Permafrost) and washed with purified water. The collagen hybridizing peptide (CHP), Cy3
conjugate dye was purchased from 3Helix. Two of the three sections on the slides were stained
with CHP while the third was incubated with PBS as a control. For CHP staining, a solution of 10
μM CHP in PBS was heated at 80 oC for 5 minutes and then quenched in an ice water bath and
centrifuged. 20-50 μL of the CHP solution (or enough to completely cover the sections) was
incubated in a refirdgerator overnight over tissue sections. After incubating with CHP, the samples
were washed three times in PBS and then mounted using a DAPI-conatining mounting medium.
Slides were imaged using a Zeiss Axiovert fluorescence microscope at 20x and 63x magnifications
using the tetramethylrhodamine (TRITC) and DAPI filter. For SHG microscopy, slides were
imaged using an Olympus FV1000 MPE Multiphoton Laser Scanning Confocal Microscope at the
Campus Microscopy and Imaging Facility (CMIF) at OSU.
6
Statistical analysis: Data are presented as means ± standard deviations. Statistically significant
differences in D-periodicity depth were determined using ANOVA analysis and Tukey-Krammer
test performed using SAS JMP software (version 14.0). The data for each of these tests complied
with the constraints of the test as validated by normality and equal variance tests. A p-value
<0.05 was considered significant.
Results and Discussion
The collagen fibrils are formed by a well-defined staggered arrangement and organization
of collagen monomers within the fibril resulting in a natural periodic distribution of gap and
overlap regions throughout its length. This molecular packing gives rise to the collagen’s
characteristic D-periodic banding. Normal collagen fibrils can thus be easily identified by their
well-defined D-periodicity and straight contours. A departure from these characteristic features
can help identify collagen fibril abnormalities.
Atomic force microscopy was utilized to evaluate ultrastructural difference in the collagen
fibril structure between healthy and AAA tissue. Normal collagen was easily identified in both
saline-infused mice and in AngII-infused mice (Figure 1, A and B respectively). Similarly, normal
collagen was recognized in both control human samples and AAA clinical human tissue (Figure
1, D and E respectively). However, both murine AAA and human AAA tissues also revealed a
population of fibrils which exhibited structural abnormalities (Figure 1, C and F respectively and
Figure 2). The abnormal collagen fibrils were characterized by D-period bands, which were
compromised or unresolvable. The fibril contours were more difficult to identify as compared to
normal fibrils.
7
Figure 1: Normal and abnormal collagen fibrils characterized in (A-C) mouse and (D-F) human AAA using atomic force microscopy (AFM). Compromised or unresolvable D-bands were observed in AAA. Arrows (in C) indicate location and direction of normal (black) or abnormal (white) collagen. Human AAA consisted of regions with both normal and abnormal collagen fibrils. All scale bars = 200nm.
8
Figure 2: Additional examples of abnormal collagen fibrils in AngII-infused mice and human AAA. AFM amplitude images show adventitial collagen fibrils with disrupted or unresolvable D-bands and display collagen fibril abnormalities across multiple samples of AngII-infused mice (A-C) and clinical human AAA (D-F). All scale bars 200nm.
Since AFM can track nanoscale topography, we utilized AFM height images and their
corresponding sectional profiles to analyze D-depth differences between normal and abnormal
collagen fibrils. The D-period depth of three groupings of collagen was identified: normal collagen
in saline-infused mice (Figure 3, A-B), normal collagen in AngII-infused mice (Figure 3, C-D)
and abnormal collagen in AngII-infused mice (Figure 3, E-F). Distributions of the D-period depth
indicate that normal collagen fibrils show no statistical significance in either AngII-infused mice
or saline-infused mice (Tukey-Kramer test, p=0.973). However, abnormal collagen fibrils found
in AngII-treated mice exhibited decreased D-depths (one way ANOVA followed by Tukey-Kramer
9
test, p˂0.0001). In some cases, D-period bands were compromised to such an extent that D-depth
could not be identified.
AFM height images were also used to analyze differences in D-period depth in normal and
abnormal collagen in human tissue. Similarly to the mouse model, D-period depth of three groups
of collagen was identified: normal collagen in clinical control human tissue (Figure 4, A-B),
normal collagen in clinical AAA tissue (Figure 4, C-D) and abnormal collagen in clinical AAA
tissue (Figure 4, E-F). As in the mouse model, normal collagen in control human tissue and normal
collagen in human AAA tissue do not present any statistically significant difference in D-period
depth. However, abnormal collagen fibrils found in AAA exhibited decreased D-depths (one way
ANOVA followed by Tukey-Kramer test, p˂0.0001). From this data, it can be elucidated that a
reduced D-depth is characteristic of abnormal collagen in AAA.
10
Figure 3: Characterization of depth of D-periods in collagen fibrils from murine aorta. AFM height images of normal fibrils and their corresponding section profiles for adventitial collagen in AngII-(C,D) and saline-infused mice (A,B) reveal a similar depth of D-period. However, abnormal collagen fibrils found in AngII-treated mice exhibited decreased D-depths (E,F). Scale bars are 200nm. Histogram shows the distribution of D-period depth from a representative mouse sample as indicated (G). Average depth of D-periods from all samples shows no significant differences between normal fibrils from each treatment group. However significantly reduced D-depth were observed for abnormal fibrils in AngII-infused mice (*p˂0.0001) (H).
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Figure 4: Characterization of depth of D-periods in collagen fibrils in human AAA. AFM height images of collagen fibrils and their corresponding section profiles are shown for normal fibrils from human control (A,B) and clinical AAA normal (C,D) and for abnormal fibrils in clinical AAA (E,F). Scale bars are 200 nm. Histogram shows the distribution of D-period depth from are presentative human sample as indicated (G). Average depth of D-periods from all samples shows no significant differences between normal fibrils from control or AAA groups. However, significantly reduced D-depth were observed for abnormal fibrils in AAA (*p˂0.0001) (H). Results from the D-depth analysis for all conditions (normal collagen in saline-infused
mice, normal collagen in AngII-infused mice, abnormal collagen in AngII-infused mice, normal
collagen in control human tissue, normal collagen in AAA human tissue and abnormal collagen in
AAA human tissue) is summarized quantitatively in Table 1. The data presented in Table 1
12
reiterates that there is no difference between normal collagen in control and AAA samples for both
murine and human AAA models. Abnormal collagen, however, has a significantly smaller D-depth
for murine and human AAA tissues. Overall, normal collagen fibrils in mouse tissue have a smaller
D-depth than collagen fibrils in human tissue. Despite the differences in D-depth of normal
collagen of mouse and human tissues, D-depth of abnormal human and murine collagen fibrils
decrease to a similar value (~1.0 nm).
Table 1: Summary of D-depth measurements of collagen fibrils in murine/human samples.
Further analysis of collagen D-periodicity revealed that although banding was
compromised in or unresolvable in abnormal collagen, there was no difference in the length of D-
periods (66 ± 6 nm, one way ANOVA, p=0.473) between abnormal and normal collagen for both
human and murine tissues.
Thus our AFM analysis coupled with electron microscopy analysis performed by others19
confirmed the presence of collagen fibril structure abnormalities in AAA in both human and mouse
AAA. However, the source of the formation of these abnormalities is not yet understood.
13
Implications of these abnormalities on the functional properties of the vessel wall and stability of
an aneurysm are also not yet well understood. Identifying the progression and development of
abnormal collagen in AAA can lead to a better understanding of how to diagnose and treat many
vascular diseases.
As another test to determine abnormal collagen we used Collagen hybridizing peptide
CHP, a reagent which stains degraded collagen. CHP binds to unfolded collagen chains, and
therefore is used to locate molecular denaturation of collagen and provides information on
structural damage of collagen. Fluorescent imaging of CHP stained murine and human tissue was
done to determine locations of degraded collagen. CHP location is represented by fluorescent
regions (Figure 5, B-C, E-F). Saline-infused murine tissue (Figure 5, A) and clinical control human
tissue (Figure 5, D) demonstrate no regions of intense fluorescence. However, AngII-infused
murine tissue (Figure 5, B-C) and human AAA tissue (Figure 5, E-F) present with several regions
of intense fluorescence. CHP analysis of aortic tissues indicates that denatured collagen is present
in AAA tissue but is not in healthy tissue. Fluorescent imaging of CHP stained aortic tissue shows
that collagen is denatured during the remodeling process of AAA.
14
Figure 5: Normal and abnormal aorta tissue is observed for collagen degradation in mouse samples (A-C) samples. Collagen hybridizing peptide (CHP) staining was done for both saline-infused mice (A) and Ang-II infused mine (B and C) and for clinical control human tissue (D) and clinical AAA human tissue (E-F). CHP binds to unraveled or degraded collagen and can therefore be used to locate degraded collagen. Saline-infused murine tissue show no degraded collagen regions. AAA tissues present regions of degraded collagen marked with fluorescent regions indicated with arrows (B and C). Second harmonic generation microscopy is a laser imaging technique used to visualize
molecular structures of tissues. Normal collagen type I and III, which is the collagen that is found
in vascular tissue ECM, naturally produces SHG contrast due to its molecular structure. On the
other hand, as previously mentioned, CHP binds to denatured collagen. By simultaneously using
these two detection methods, the locations of degraded and healthy collagen were able to be
located. The red signal represents degraded collagen (CHP signal), the green represents healthy
collagen (SHG signal) and blue represents cell nuclei (DAPI signal). The saline-infused control
murine sample (Figure 6, A) demonstrated that healthy aortic tissue lacks any degraded collagen
but shows evidence of healthy collagen. However, AAA murine (Figure 6, B) tissue and AAA
human tissue (Figure 6, C-E) provide evidence of degraded collagen. Denatured collagen and
15
healthy collagen locations are clearly identifiable by their distinct fluorescent colors. Abnormal
collagen regions are not co-localized with healthy collagen regions, but rather are isolated from
one another. Additionally, the SHG images provide qualitative and quantitative information about
the state of cell morphology and integrity. Cell nuclei are evident both in regions containing normal
and abnormal collagen indicating that collagen degradation is not a result of cell death. Instead it
is attributed to the collagen remodeling process of AAA.
Figure 6: Images showing SHG and CHP signal. Mouse (A-B) and human (C-E) tissue is observed to determine location of degraded and heathy collagen. Blue fluorescence represents DAPI staining, red fluorescence represents CHP signal and green fluorescence show SHG signal. Healthy tissue possesses no degraded collagen (A). SHG and CHP signals demonstrate the presence of degraded collagen in murine (B) and human (C-E) AAA tissue.
SHG analysis confirmed information identified with fluorescent imaging of CHP stained
tissues. Denatured collagen occurs as a result of the remodeling process associated with AAA, and
is localized to sporadic pockets throughout the remodeled region of tissue. Quantification of the
CHP and SHG signal can provide an estimate of the percent of abnormal and normal collagen in
AAA. Understanding more about the extent of collagen degradation in AAA progression can
provide insights into the collagen remodeling process.
16
Conclusion
In summary, abnormal collagen fibrils found in AAA present distinct differences from
normal collagen found both in control samples and in AAA samples. AFM analysis demonstrated
that abnormal fibrils have compromised D-period bands, undulating appearance and decreased
organization. While abnormal collagen was only found in AAA tissue, normal collagen was found
in both control and AAA samples. This indicated that abnormal collagen is characteristic of
collagen remodeling in AAA. Quantitative analysis on compromised collagen D-period bands
provided information about the differences in D-depth in normal and abnormal collagen. Normal
collagen found in saline- and AngII-infused mice and in clinical control and AAA human tissue
are not significantly different. Abnormal collagen fibrils, however, have a decreased D-depth
present in both murine and human tissue indicating that diminished D-depth is another trait of
AAA. Additionally, fluorescent imaging and SHG analysis of CHP stained aortic tissues revealed
that denatured collagen is present in AAA tissues but not in control tissues. Denatured collagen
and healthy collagen are located in different regions along tissue samples. Cell nuclei are located
in both regions indicating that collagen degrades as a result of the remodeling process in AAA and
not because of tissue death. Quantification of SHG and CHP signals in fluorescent images can
provide more information about the progression of collagen remodeling in AAA. Moving forward,
a technique will be optimized to do so which can provide an estimate of the percent of abnormal
and normal collagen. Additionally, adjacent sections of human and murine tissue will be analyzed
for AFM and CHP signal simultaneously to determine if and the type of collagen abnormalities
are present in areas populated with collagen degradation. Understanding more about the
remodeling process can provide novel insights into the development of diagnostics and
therapeutics for AAA and other vascular diseases.
17
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